A dual organelle-targeting mechanosensitive probe

Cells are responsive to the mechanical environment, but the methods to detect simultaneously how different organelles react in mechanobiological processes remain largely unexplored. We herein report a dual organelle-targeting fluorescent probe, (E)-1-[3-(diethoxyphosphoryl)propyl]-4-[4-(diethylamino)styryl]pyridin-1-ium bromide (ASP-PE), for mechanical mapping in live cells. ASP-PE is aggregation-induced emission active and is sensitive to the local mechanical environment. It targets the plasma membrane (PM) and intracellular mitochondria in cells by its phosphonate moiety and pyridinium. In this work, through ASP-PE staining, changes of membrane tension in the PM and mitochondria in response to varied osmotic pressure and substrate stiffness are visualized using fluorescence lifetime imaging microscopy. The mechanobiological importance of actin filaments and microtubules in the PM and mitochondria is also investigated using this probe. Computational simulations are applied to study the sensing mechanism of the probe. This study introduces a unique tool for mapping the membrane tension in the PM and mitochondria together, providing us great opportunities to study organelle’s interactions in mechanobiology.


INTRODUCTION
Mechanical forces exerted by the extracellular environment are counterbalanced by intracellular pressure and play important roles in cellular physiology and homeostasis. The impacts of these biophysical stimuli are evident in dictating cell fate and behaviors, and mechanical cues in the cellular microenvironment have been found important in many biological processes such as cancer progression (1). For example, mechanical fluctuations in the cellular microenvironment, including tensile stress (2), hydrostatic pressure (3), shear stress (4), interfacial geometry (5), and matrix stiffness (6), are shown influencing and governing carcinogenesis (1,7). Understanding the relationships between mechanobiology and cancers can thus lead to the development of new diagnostic methods and therapeutical interventions. Micropipette aspiration and atomic force microscope cantilevers (or optical/magnetic tweezers) are the most commonly used techniques for evaluating cell mechanical properties or imaging mechanobiological events (8). These methods can provide quantitative measurements. However, they require experienced operators. Moreover, only the mechanical property of a single target can be assessed in each single measurement, and the intracellular compartments are not directly accessible for these contact-based methods. Therefore, convenient methods rendering spatialtemporal information of mechanical properties in multiple organelles are highly desirable in the field of mechanobiology to complement the current tools and to provide information that is inaccessible using the current methods.
The plasma membrane (PM) is the foremost and pivotal interface that mediates cell mechanobiological responses to external mechanical stimuli (9). For example, an environment of high stiffness induces the metastatic phenotype mechanotransduced by integrin (6,10). To better understand the mechanobiology in live cells, several fluorescence imaging techniques focusing on the PM have been exploited, such as fluorescence recovery after photobleaching, fluorescence resonance energy transfer, and Laurdan imaging (11)(12)(13). In addition to the PM, other intracellular organelles, especially mitochondria, are also important targets in mechanobiological studies. In particular, how mitochondrial dynamics and biogenesis are modulated in response to mechanical stimuli during epithelialmesenchymal transition and metastasis have been intensively studied (4,(14)(15)(16)(17).
Recently, Matile et al. (18)(19)(20)(21) have reported a series of dithienothiophene-based fluorescent membrane tension (σ) probes that locally sense one designated organelle membrane each time in live cells. Although mechanical stress can be transmitted from the PM to intracellular organelles, tools to unveil simultaneously how the PM and intracellular organelles respond to mechanical stress are highly desirable but not yet available. Along with the development of conventional bioprobes, fluorescent probes that exhibit aggregationinduced emission (AIE) have emerged rapidly in the field of bioimaging. AIE-active fluorophores emit weakly or are even nonemissive in molecularly dissolved state with active intramolecular motions, but they emit intensively when the intramolecular motion is restricted (22)(23)(24). This mechanism, named restriction of intramolecular motion (RIM), has been used to develop various fluorescent probes (25).
Here, we designed and synthesized a mechanosensitive dual PMmitochondria-targeting AIE-active fluorescent probe named ASP-PE. Unlike the existing tools that track only the PM but no other intracellular compartments simultaneously, ASP-PE is a bioprobe that specifically targets both the PM and mitochondria at the same time. We demonstrated that ASP-PE could detect single-cell mechanotype from its fluorescence lifetime (τ f ) in the PM and mitochondria under external mechanical stimuli including tension, compression, and changes in substrate stiffness. Distinct mechanotype changes in the PM and mitochondria under different mechanical disturbances were revealed by ASP-PE. The role of cytoskeletons in cellular mechanobiology was also demonstrated using ASP-PE. Molecular dynamics (MD) investigation coupled to a stimulated phospholipid bilayer structure was used to investigate the relationship between membrane tension and fluorescence lifetime of the fluorescent probes in silico. Structural comparisons among ASP-PE and two styryl dyes suggested that the phosphonate moiety of ASP-PE was critical for its PM targetability and high mechanosensitivity in the target membrane.

Molecular design and photophysical properties of ASP-PE
The new mechanosensitive probe ASP-PE (molecular structure shown in Fig. 1A) was synthesized from commercially available chemical reagents through a straightforward two-step synthetic route as shown in fig. S1 (see the synthetic details in the Supplementary Materials). ASP-PE is constituted of an electron push-pull πconjugation skeleton. It is soluble in common organic solvents such as dichloromethane, methanol, ethanol, and dimethyl sulfoxide (DMSO) but insoluble in most hydrophobic solvents, such as hexane, diethyl ether, and ethyl acetate (EA). The intermediate (E)-N,N-diethyl-4-(2-[ pyridin-4-yl)vinyl]aniline and ASP-PE were structurally characterized by 1 H-and 13 C-nuclear magnetic resonance (NMR) spectroscopy as well as high-resolution mass spectrometry (HRMS) (figs. S2 to S4). The photophysical properties of ASP-PE were studied and illustrated in Fig. 1. ASP-PE exhibited an absorption peak at the wavelength (λ abs ) of 500 nm with a molar absorptivity (ε) of ca. 5 × 10 4 M −1 cm −1 and an emission peak (λ em ) at ca. 610 nm in ethanol solution (10 μM; photoexcitation at 515 nm) (Fig. 1B). Its absorption and emission spectra in different single solvents are given in fig. S5. When the EA fraction of the solution(s) increased beyond 0.7, ASP-PE tended to reach the aggregated state due to the lower solvation power of solvent mixtures. An enhanced photoluminescence (PL) signal with a minor and hypsochromic shift (ca. 10 nm) could be found in the solutions with aggregates [i.e., EA:EtOH = 8:2 or 9:1 (v/v)], demonstrating an AIE characteristic (Fig. 1C). In a viscous environment with a high glycerol (GLY) fraction, the probe exhibited stronger fluorescence and longer fluorescence lifetime under confinement (Fig. 1, D and E).

Biocompatibility and dualorganelle targetability of ASP-PE in live cells
The cytotoxicity of ASP-PE was evaluated using the Cell Counting Kit-8 (CCK-8) assay before bioapplication. As illustrated in fig. S6, the cell viability was more than 70% at a concentration below 15 μM, demonstrating a good biocompatibility of ASP-PE in vitro. We then tested its performance in live-cell imaging by incubating HeLa cells with 10 μM ASP-PE for 15 min. ASP-PE specifically labeled the cell surface and intracellular compartments. Costaining experiments showed that these signals were colocalized with the commercial PM and the mitochondria probes (CellMask Deep Red Plasma Membrane Stain and BioTracker 405 Blue Mitochondria Dye, respectively) ( Fig. 2A). Overall,~80% of the ASP-PE signal overlaid with the PM and mitochondria (Fig. 2B), revealing that the dual targetability of ASP-PE to the PM and mitochondria was highly selective.
To understand the chemical structure of ASP-PE that attributes to this dual targeting specificity, we experimentally compared ASP-PE with two structurally similar styryl dyes (structures shown in Fig. 2A). The first one is (E)-4-[4-(diethylamino)styryl]-1-methylpyridin-1-ium iodide (ASP) with one methyl group locking to the pyridinium ring, which is reported as a mitochondrial probe for live-cell imaging (26). The second one is (E)-4-[4-(diethylamino)styryl]-1-(2-octyldodecyl)pyridin-1-ium bromide (ASP-OD) with a long and branched alkyl chain on the pyridinium, which was structurally characterized by 1 H-and 13 C-NMR spectroscopy as well as HRMS (figs. S7 to S9). The photophysical properties of ASP and ASP-OD were depicted in the ultraviolet-visible (UV-Vis) absorption spectra (fig. S10), PL spectra (figs. S10 and S11), and time-correlated single-photon counting (TCSPC) spectra (fig. S12). These data showed that all three molecules have similar photophysical properties because of the same π-conjugation moiety. In the fluorescence imaging, ASP preferentially labeled the mitochondria in HeLa cells ( Fig. 2A). With the long and branched side chain, ASP-OD targeted to the mitochondria and weakly outlined the cell boundary ( Fig. 2A). By comparing the subcellular distribution, ASP-PE exhibited the best PM and mitochondria localization among the above three molecules (Fig. 2, B and C). We inititally postulated that the electronegative diethyl phosphonate moiety in ASP-PE mimics the negatively charged phospholipid head, which provides better PM retention than the hydrophobic interactions between the branched alkyl chain in ASP-OD and the fatty acid tails of phospholipids. Meanwhile, the pyridinium in ASP-PE, as in ASP and ASP-OD, allows its mitochondrial localization. These, therefore, confer ASP-PE with dual targetability to the PM and mitochondria.
Distinct τ f of ASP-PE in PM and mitochondria and its sensitivity to osmotic pressure The average ASP-PE lifetime in the PM and mitochondria were distinguishable under fluorescence lifetime imaging microscopy (FLIM) at 4.73 and 3.64 ns, respectively (Fig. 3). In comparison, the mean τ f of ASP and ASP-OD, which did not have the (diethoxyphosphoryl)propyl moiety, was measured in cells (figs. S13 and S14). The mean τ f of ASP in the PM and mitochondria was 4.19 and 2.65 ns, respectively, whereas the mean τ f of ASP-OD in the PM and mitochondria was 1.55 and 1.28 ns, respectively. Their τ f ranges or mean τ f differences in the PM and mitochondria were not substantial when comparing to ASP-PE.
It is reported that osmotic challenge induced by the hypotonic condition, in which the osmolarity was lower than iso-osmolarity (360 mOsm/liter), causes tensile stress, whereas the hypertonic condition causes compressive pressure (27). To determine whether ASP-PE lifetime is sensitive to changes in mechanical properties, osmotic shock was applied to ASP-PE-stained cells. By using FLIM, ASP-PE showed opposite fluorescence lifetime changes in HeLa cells challenged with hypotonic (75 mOsm/liter) or hypertonic (750 mOsm/liter) solutions when comparing with the isotonic control (Fig. 4A). The mean τ f of ASP-PE in the PM increased significantly under hypotonic shock, but it decreased upon hypertonic treatment (Fig. 4, B and C). The mean τ f of ASP-PE in the PM was inversely proportional to the osmolarity of the incubation buffer with the R 2 (coefficient of determination) of 0.9858 (Fig. 4D). The linear correlation of the τ f of ASP-PE in the PM under varied osmotic stress indicates a primary mechanobiological change in the PM, which is possibly caused by altered membrane metrics such as membrane tension and lipid phase separation (18,28,29). Moreover, we also observed a similar phenomenon for the mean τ f of ASP-PE in mitochondria under both hypoosmotic and hyperosmotic conditions as compared with the isotonic condition ( Fig. 4, A, E, and F). A linear correlation of the mean τ f of ASP-PE as a function of osmolarity was found with the R 2 of 0.9898 from 162 to 750 mOsm/liter (Fig. 4G). Regarding to ASP (fig. S13) and ASP-OD (fig. S14), their fluorescence lifetimes were less sensitive to osmotic shock. So, the floppy moiety of diethyl phosphonate in ASP-PE acts like a "mechanosensitivity enhancer" toward the membranous organelles without alternating the π-conjugation skeleton of the fluorescent unit. As a result, ASP-PE can be used to sense changes of mechanical properties in the PM and mitochondria.

Varied substrate stiffness alters PM tension
The cellular microenvironment in vivo is complex and dynamic with stiffness heterogeneity that induces cellular adaptation (1,6). For example, the metastatic behavior of mammary cancer cells can be influenced by matrix stiffness (30). The importance of substratedependent mechanobiological response prompts us to further examine whether ASP-PE can detect these influences with varied substrate stiffness.
To fabricate substrates with softer stiffnesses compared to the glass, commercially available polydimethylsiloxane (PDMS; SYLGARD 184) was used. Its mechanical property was revealed by its stress-strain curve where Young's moduli was 855 kPa for the PDMS. In addition, the thickness of these PDMS thin films was controlled to be around 90 μm. It was thicker than the critical value of 38 μm which is the marginal thickness for cells responding to substrate stiffness (31). Cells were seeded on the PDMS surface and a glass-bottom culture dish, respectively, for comparison. Under FLIM, HeLa cells seeded on PDMS substrates showed a rounder morphology, and their lifetime maps of ASP-PE were different to those seeded on the glass surface with a Young's modulus in gigapascals (Fig. 5A). For cells grown on the PDMS substrate, the mean τ f of ASP-PE in the PM decreased significantly compared with cells grown on the glass surface (Fig. 5, B and C). On the other hand, the mean τ f of ASP-PE in mitochondria showed no significant difference among them (Fig. 5, D and E). Provided that adherent cells can sense and respond to substrate stiffness so that stiffer substrates result in more rigid cells with higher tension overall (32), our data are consistent with that and demonstrate that ASP-PE lifetime can sense changes in PM tension upon varied substrate stiffness from kilopascals to gigapascals.

The relation of the cytoskeleton and membrane tension is revealed by ASP-PE
The cytoskeleton is a highly dynamic protein network that provides cells with structural support, a framework for organelle movement, and a system for generation of mechanical forces. The fence and picket model proposes that actin filaments extensively tether to the PM that confines the lateral movement of molecules in the membrane (9,33). This lateral organization structurally strengthens the PM against external forces. On the other hand, microtubules are involved in maintenance of cell shape, cell adhesion, as well as organelle positioning and trafficking.
To study the importance of the cytoskeleton in mechanobiology, we destabilized microtubules and actin filaments by nocodazole (Noc) and cytochalasin D (CytoD), respectively (34,35). Fluorescence imaging showed that microtubules were fragmented by Noc ( fig. S15A), while networks of actin filaments were disassembled by CytoD ( fig. S15B). We then investigated the τ f difference of ASP-PE in the PM and mitochondria under the pharmacological treatments. Under FLIM, disorganization of microtubules decreased the mean τ f of ASP-PE in the PM of Noc-treated cells (Fig. 6, A to C) but not significantly changed ASP-PE lifetime in mitochondria (Fig. 6, D and E). This implies that cell contraction due to microtubule disruption decreases the PM tension. Conversely, disassembly of actin filaments had no significant effect on the mean τ f of ASP-PE in the PM of CytoD-treated cells (Fig. 6, F to H) but decreased ASP-PE lifetime in mitochondria (Fig. 6, I and J). This suggests that cell shrinkage following actin depolymerization exerted increased intracellular stress on mitochondrial membrane, leading to a decreased membrane tension in the mitochondrial membrane.

MD simulation investigation MD simulations of probe-membrane interactions
To gain further insight into the probing mechanism, MD simulation was used to unravel the relationship between the structures of staining agents and their mechanical sensitivities toward the environment of phospholipid bilayers. Before determining the free-  (Fig. 7A). On the basis of the equalized phospholipid membrane, three phospholipid membrane models with their phospholipid membrane centers occupied by the ASP, ASP-PE, and ASP-OD molecules, respectively, were built (fig. S16). Last, the pulling MD simulations were used to pull the three molecules from the center of the phospholipid membranes to the water phase ( fig. S17), and the resultant simulation windows along the pulling paths were further used to perform the bias MD simulation with umbrella sampling to sampling. The overlap degree between the histogram distributions of any two adjacent windows was used to evaluate the adequacy of sampling ( fig. S18).
On the basis of the sampled data from all simulation windows and the weighted histogram analysis method techniques (36), the relative free-energy profiles of translocation of ASP, ASP-PE, and ASP-OD molecules from the center of the phospholipid membrane to the water phase were presented in Fig. 7B. The phospholipid membrane center and the water phase served as the reference of Z position as 0 Å and free-energy shifted to 0 kcal/mol, respectively. For the entry of ASP and ASP-PE molecules to the phospholipid membrane, their free-energy profiles dropped to −2.51 and −4.50 kcal/mol, respectively, after an almost negligible energy barrier and reached to the global minimum state. However, comparing to the Z position of ASP molecule (Z = 18 Å) at its global minimum, the Z position of ASP-PE molecule (Z = 11 Å) was closer to the phospholipid membrane center. From the global minimum to the global maximum in the phospholipid membrane center (Z = 0 Å), both the free-energy profiles of ASP and ASP-PE molecule needed to undergo a continuous endothermic process to overcome the energy barrier. For the entry of ASP-OD molecule to the phospholipid membrane, its free-energy profile first dropped to −16.20 kcal/ mol and reached a minimum at Z = 12 Å. After overcoming an almost negligible energy barrier, its free-energy profile further dropped sharply until it reached the global minimum state in the phospholipid membrane center (Z = 0 Å) whose free-energy was −26.82 kcal/mol. From the simulation, it is shown that although these three molecules have the same π-conjugation moiety, they present different stable positions (i.e., global minimum) and translocation abilities (i.e., the energy barrier that had to overcome from the water phase to the phospholipid membrane center) in the phospholipid membrane. The calculated effective resistance coefficients and effective permeability coefficients are shown in table S1.
To unravel the intrinsic mechanism for the differences of the most stable positions and translocation abilities of ASP, ASP-PE, and ASP-OD molecules, interactions between these three molecules and the phospholipid membrane in their most stable positions were analyzed. As shown in Fig. 7C, when the ASP molecule was in the most stable position (Z = 18 Å), it inserted vertically into the phospholipid membrane with its positively charged pyridine ring surrounded by the negatively charged phosphatidylcholine/ sphingomyelin head groups. This insertion mode could enhance the electrostatic interaction between these two groups. However, when the ASP-PE molecule was in the most stable position (Z = 11 Å), it laid parallel to the phospholipid membrane with its positively charged pyridine ring was in proximity to but not surrounded by the negatively charged phosphatidylcholine/sphingomyelin head groups (shown in Fig. 7C). This parallel orientation could be attributed to the diethyl phosphate group next to the pyridine ring, which, to a certain extent, played a role in electrostatic repulsion between the ASP-PE molecule and phosphatidylcholine/sphingomyelin head groups.
The phospholipid membrane center was the most stable position (Z = 0 Å) of the ASP-OD molecule (shown in Fig. 7C). This arrangement may be related to the presence of hydrophobic n-octyl and ndecyl chains next to the pyridine ring. It enhanced the hydrophobic interaction between the ASP-OD molecule and the alkyl chains in the phospholipid membrane, which allows complete anchoring of the ASP-OD molecule. This postulation is consistent with the negatively charged phosphatidylcholine/sphingomyelin head groups that are in proximity to the positively charged pyridine ring of the ASP-OD molecule. From the above analysis, we concluded that it was the structural differences of the photo-inactive moiety among ASP, ASP-PE, and ASP-OD molecules that determined their most stable positions and postures in the phospholipid membrane.

Fluorescence properties of ASP, ASP-PE, and ASP-OD
Besides the most stable positions and translocation abilities, fluorescence properties of ASP, ASP-PE, and ASP-OD molecules were also simulated and calculated. On the basis of the conformation of ASP, ASP-PE, and ASP-OD molecules in the pure ethanol solvent and the phospholipid membrane randomly extracted from umbrella sampling structures at the window with the largest partition (i.e., the most stable position; fig. S19), their structural changes, electronic excitation properties, and reorganization energies were used to determine the fluorescence properties. Notably, different membrane tensions (+50, 0, and −200 dyne/cm) were considered for each probe conformation in the phospholipid membrane during the quantitative calculation of fluorescence properties.
First, the structural changes between the ground state (S 0 state) and the first excited singlet state (S 1 state) were considered. As shown in fig. S20, the root mean square deviations (RMSD; an indicating value showing the degree of similarity or superimposition of conformations) between S 0 state and S 1 state for the ASP, ASP-PE, and ASP-OD molecules in the pure ethanol solvent were 0.067,  0.111, and 0.138 Å, respectively. When these molecules were embedded in the phospholipid membrane with a surface tension of +50 dyne/cm, the RMSDs for the ASP, ASP-PE, and ASP-OD molecules were 0.049, 0.088, and 0.087 Å, respectively. When the surface tension of phospholipid membrane was changed to 0 dyne/cm, the corresponding RMSD values decreased to 0.037, 0.067, and 0.065 Å, respectively. When the surface tension of phospholipid membrane was −200 dyne/cm, they further decreased to 0.030, 0.024, and 0.037 Å, respectively. Obviously, the phospholipid membrane could reduce the conformational fluctuations of ASP, ASP-PE, and ASP-OD molecules and increase their rigidity. The more order the phospholipid membrane is, the smaller fluctuations and more rigid of the embedded dyes will be ( fig. S21).
Second, the reorganization energies (RE) of S 0 state and S 1 state were further considered. As shown in fig. S20, when the ASP, ASP-PE, and ASP-OD molecules translocate from the pure ethanol solvent to phospholipid membranes, all their RE decreased from 21 to 24 kcal/mol to 4 to 6 kcal/mol. Moreover, the more ordered phospholipid membrane resulted in the lower RE of the embedded ASP, ASP-PE, and ASP-OD molecules. Combining the results of structural changes and reorganization energies, it was shown that the ASP, ASP-PE, and ASP-OD molecules in the pure ethanol solvent underwent relatively large structural changes upon photoexcitation because of the high freedom of intramolecular motion. This leads to high RE to recover the structural changes caused by the molecular motion. However, when the dye molecules were embedded in the phospholipid membrane, their freedom of molecular motion naturally decreases due to the densely packing nature of the phospholipid membrane. Therefore, during the electron excitation from S 0 to S 1 state, the ASP, ASP-PE, and ASP-OD molecule only underwent relatively small structural motions and with lower RE to recover the conformations of the ground state.
Considering that the recovery of structural rotations primarily goes through the nonradiative transition, the nonradiative decay rate constants of membrane-embedded ASP, ASP-PE, and ASP-OD molecules will decrease due to the limitation of motion. This was confirmed by the calculated nonradiative decay rate constant for ASP-PE molecule as shown in Table 1 and table S2. Meanwhile, the decrease of nonradiative decay rate constant upon the increased membrane tension causes the higher quantum yield and longer fluorescence lifetime.
For ASP-PE molecule, when the surface tension adopted on the phospholipid membrane increased, the limitation on intramolecular motion led to a decreased nonradiative rate constant (which was The phosphatidylcholine head groups of POPC phospholipids and the sphingomyelin head groups of PSM phospholipids were shown as orange spheres. The palmitoyl acyl chains and oleoyl acyl chains of POPC phospholipids were shown as cyan sticks and limon sticks, respectively. The palmitoyl acyl chains and sphingosine acyl chains of PSM phospholipids were shown as cyan sticks and pink sticks, respectively. The CHOL phospholipids were shown as light blue sticks. The ASP, ASP-PE, and ASP-OD molecules were shown as green spheres. Meanwhile, the neutralizing K + ions and Cl − ions were shown as purple spheres and magenta spheres, respectively. The water molecules were shown as red spheres. also understood as a RIM effect), this increases the quantum yield and fluorescence lifetime overall. Compared to the RMSD and RE values of ASP and ASP-OD, those of ASP-PE showed a higher sensitivity to the membrane tension changes, indicating that APE-PE is the best membrane tension probe among the three molecules here, which is in good agreement with the experimental results. Considering that ASP, ASP-PE, and ASP-OD molecules share the same chromophore unit, their photo-inactive moieties led to their most stable positions and postures in the membrane, which, in turn, led to their different sensitivities to the changes of membrane tension.

DISCUSSION
In summary, with an electron push-pull π-conjugation skeleton and a rarely reported diethyl phosphonate moiety on the pyridinium ring, an AIE-based bioprobe ASP-PE is developed for dual PM-mitochondria-targeting and visualizing cellular mechanical properties. Referring to respective subcellular localization and biomechanical sensitivity of ASP-PE and two structurally similar styryl dyes, the structural comparisons suggest that the phosphonate ester moiety in ASP-PE acts as a mechanosensitivity enhancer in membranous organelles. In addition to sensitivity to different environments during livecell imaging by FLIM, two distinguishable fluorescence lifetimes are exhibited in the PM and mitochondria, respectively, for ASP-PE.
In high-tension membrane, ASP-PE experiences less freedom of molecular motions and exhibits brighter emission and longer fluorescence lifetime due to the RIM effect. The fluorescence lifetime of ASP-PE in the cell membrane of HeLa cells is linearly correlated with the osmotic pressure in the medium. With the dual targeting feature of ASP-PE, changes of mechanical properties not only in the PM but also in intracellular mitochondria are studied. Different mechanobiological responses in these membrane systems are revealed by the τ f changes of ASP-PE under osmotic shock and varied substrate stiffness. In hypotonic medium or on the high stiffness substrate, the cell membrane is stretched and has a high membrane tension. The probe correspondingly shows a significantly longer fluorescence lifetime. In hypertonic medium or on the low stiffness substrate, the fluorescence lifetimes of ASP-PE in the cell membrane are much shorter. The signal of ASP-PE indicates that the mechanical properties of mitochondria are almost unaffected by substrate stiffness as the PM does. While under osmotic stress, the mitochondria have altered fluorescence lifetime signals with the same trend as the PM. It demonstrates that the probe can be used in studying the relation of the PM and mitochondria.
Furthermore, ASP-PE staining under pharmacological treatments reveals that the PM and mitochondrial membranes are distinctively influenced by microtubules or actin disorganization. This further implies that PM and mitochondrial membranes encounter respective challenges during mechanical stress. Meanwhile, it should be noted that cell contraction or cell volume changes accompanied may alter the focal planes of the PM and mitochondria differently in adherent cells. In those scenarios, it may be challenging to unmix the signals from the PM and mitochondria under the wide-field FLIM. In addition to the experimental data, in silico investigation is used to understand and verify the hypothesis that correlates the membrane tension of PM (or mitochondria) and the fluorescence lifetime of the dual organelle targeting mechanosensitive probe ASP-PE. The computational analysis reveals that the nonπ-conjugated units present in the pyridine ring in these three molecules greatly affect their most favorite positions and binding postures in the membrane, which further led to different sensitivities to membrane tension changes.
Together, the promising sensitivity of ASP-PE to mechanical stress and mechanobiological events demonstrates a rapid and reliable approach for exploring cellular mechanosensing. Its potential application in mechanobiological studies can further be widened by modifying this mechanosensitive bioprobe with organelle-targeting signals to reveal organelle mechanosensitivity in real time during dynamic processes, such as morphogenesis and cell migration, with the aid of FLIM.

Materials and reagents
All chemical reactions were performed under an inert nitrogen atmosphere with the use of a Schlenk line. Glassware was dried in an oven before use. Commercially available reagents were used without purification. All the reagents for chemical synthesis were purchased from Tokyo Chemical Industry Co. Ltd., Sigma-Aldrich, Acros Organics, or J&K Scientific. Dry solvents were purchased from the abovementioned companies and stored in the presence of activated 3-Å molecular sieves.

Instrumentation for chemical characterizations and photophysical measurements
Proton and carbon NMR spectra were measured in CDCl 3 on a Bruker AVANCE III 500 (or 400) MHz NMR spectrometer, and tetramethylsilane was exploited as an internal standard for calibrating the chemical shift. Electrospray ionization quadrupole time-offlight (ESI-Q-TOF) MS was performed on an Agilent 6540 liquid chromatography-ESI-Q-TOF mass spectrometer. UV-Vis absorption spectroscopy was performed on a Molecular Devices Spectra-Max M2e in different solutions at 293 K. The solution emission spectra and TCSPC of the chemical probes were measured on a Per-kinElmer fluorescence spectrometer LS 55 and FLS1000 PL spectrometer at 293 K, respectively. The fluorescence decay curves were analyzed using an algorithm fitting routine supplied by Fluoracle and were shown to follow a biexponential function.  30 MHz controlled using NIS-Elements AR (Nikon, Japan) integrated with a module for the pco.flim camera. By measuring the phase shift acquired from photons emitted induced by the sinusoidally modulated laser light source, the lifetime map of the interest fields and the lifetime distributions were collected and analyzed using NIS-Elements AR (Nikon, Japan).

Fabrication of PDMS substrates
The blends of PDMS were prepared using the commercially available SYLGARD 184 (Dow Corning). A blend with 10:1 (v/v, base/ curing agent) of SYLGARD 184 PDMS was prepared by stirring. For live-cell imaging, the construction of a PDMS well-included two parts, which were a thin-bottom layer and a frame structure. All the PDMS mixtures were vacuum-degassed in a desiccator for 1 hour. Then, the thin-bottom layer was prepared by the float-onwater method (37). Briefly, we used 8 ml of ultrapure water loaded in a 60-mm petri dish as the substratum, dropped an approximately 100 μg of the PDMS blends on it using a 1000-μl pipette tip, and then left it spreading over the surface of the water. Samples were later cured at 75°C for 1 hour in a convection oven. On the other hand, the preparation of the frame structure was achieved by casting the SYLGARD 184 PDMS in a 35-mm petri dish followed by a curing process at 75°C for 1 hour. The inner circular opening (Ø = 5 mm) of the frames was created by a punch. The complete PDMS well was next assembled by putting and pressing the PDMS frame structure onto the thin PDMS layers floated on water for adhesion. The well was coated with 5 μg of type I collagen per centimeter squared at 37°C for at least 4 hours before seeding cells on it.

Cell viability
The cytotoxicity was evaluated using a standard CCK-8. Briefly, HeLa cells were seeded in a 96-well plate at a density of 7 × 10 3 cells per well and cultured overnight before treatment. A stock of 10 mM ASP-PE in DMSO was diluted to different concentrations (2.5, 5, 10, 15, or 20 μM) in fresh culture medium, and DMSO was used as the vehicle control. After incubating the cells with the above solutions for 24 hours, 10 μl of CCK-8 stock solution in PBS was added to each well, and cells were incubated for an additional 4 hours. The absorbance was measured using a SpectraMax M2e microplate reader (Molecular Devices) at a wavelength of 450 nm, and the cell viability percentage was calculated from the optical density.

Tensile testing
The tensile bar strips of the PDMS substrates were prepared by casting the PDMS blends in a mold with shapes in compliance with ASTM D412 Type C and curing at 75°C for 1 hour. Testing was done on Instron 5942 Micro Newton Tester (Instron, Norwood, MA, USA). The samples were stretched at a rate of 50 mm/min until failure, following ASTM D412. The Young's modulus of the PDMS strips was subsequently determined automatically by the in-house software provided by Instron in which the steepest slope, which is the Young's modulus, was calculated on the initial linear portion of the curve using least-square fit on the test data.

Statistical analysis
Statistical significance was determined using two-tailed unpaired t test or one-way analysis of variance (ANOVA) with Tukey's post hoc test. A P value less than 0.05 was considered statistically significant, and n is the number of independent experiments.

Computational investigation
Detailed computational methods, including (i) the preparation of ternary component-based phospholipid bilayer model, (ii) MD modeling, (iii) pulling MD simulation, (iv) free-energy profiles simulation, (v) permeability coefficient calculations, and (vi) quantum mechanics/molecular mechanics calculations are fully described in the Supplementary Materials.

Supplementary Materials
This PDF file includes: Supplementary Text Tables S1 and S2 Figs. S1 to S21 References